Antisense compounds have shown great promise as therapeutics, diagnostics and aids to therapeutic target validation. An antisense compound modulates a protein's activity by attenuating the concentration of polynucleotides, especially RNA, involved in protein synthesis. This is in contrast to conventional therapeutic methods, which seek to modulate protein activities by direct interaction between putative drugs and proteins. The effect of an antisense compound's interaction with intracellular polynucleotides is thus a predictable, albeit indirect, modulation of the activity of the protein or peptide that the cell normally manufactures using the polynucleotide as a template.
In general, antisense methods involve determining the sequence of a coding polynucleotide (e.g. mRNA) that encodes for a certain protein, developing a relatively short oligomer (antisense compound) that selectively binds to the polynucleotide (sense strand), and introducing the oligomer into the intracellular environment. Antisense methods can predictably modulate gene expression through a variety of mechanisms. In one such mechanism, the antisense strand blocks translation by competitively binding to the sense strand. In another mechanism, an antisense strand containing a stretch of DNA (e.g. phosphorothioate DNA) binds to the sense strand, and then the DNA-RNA hybrid is recognized by RNAse H, an endonuclease, which selectively cleaves the DNA-RNA hybrid, thereby reducing intracellular RNA levels. Another methodology involves the interaction between small double stranded RNA oligomers and mRNA. In such mechanisms, interaction between the RISC complex, the antisense strand of the small double-stranded RNA and intracellular mRNA results in cleavage and degradation of the mRNA.
As antisense molecules have become accepted as therapeutic and diagnostic agents, the need to produce oligonucleotides in large quantities has increased as well. The most commonly used antisense compounds to date have been oligonucleotides, phosphorothioate oligonucleotides and second generation oligonucleotides having one or more modified ribosyl sugar units, and more recently, ribosyl sugar units. The methods for making these three types of antisense oligomers are roughly similar, and include the phosphotriester method, as described by Reese, Tetrahedron 1978, 34, 3143; the phosphoramidite method, as described by Beaucage, in Methods in Molecular Biology: Protocols for Oligonucleotides and Analogs; Agrawal, ed.; Humana Press: Totowa, 1993, Vol. 20, 33–61; and the H-phosphonate method, as described by Froehler in Methods in Molecular Biology: Protocols for Oligonucleotides and Analogs Agrawal, ed.; Humana Press: Totowa, 1993, Vol. 20, 63–80. Of these three methods, the phosphoramidite method has become a defacto standard in the industry.
A typical oligonucleotide synthesis using phosphoramidite chemistry (i.e. the amidite methodology) is set forth below. First, a primer support is provided in a standard synthesizer column. The primer support is typically a solid support (supt) having a linker (link) covalently bonded thereto. It is common to purchase the primer support with a first 5′-protected nucleoside bonded thereto. 
Primer support: bg is a 5′-blocking group, Bx is a nucleobase, R2′ is H, OH, OH protected with a removable protecting group, or a 2′-substituent, such as 2′-deoxy-2′-methoxyethoxy (2′-O-MOE), and link is the covalent linking group, such as a succinyl group, which joins the nucleoside to the support, supt.                (A) The 5′-blocking group bg (e.g. 4,4′-dimethoxytrityl) is first removed (e.g. by exposing the 5′-blocked primer-support bound nucleoside to an acid), thereby producing a support-bound nucleoside of the formula:         
Activated primer support: wherein supt is the solid support, link is the linking group, Bx is a nucleobase, R2′ is H, OH, OH protected with a removable protecting group, or a 2′-substituent.                (B) The column is then washed with acetonitrile, which acts to both “push” the regent (acid) onto the column, and to wash unreacted reagent and the removed 5′-blocking group (e.g. trityl alcohol) from the column.        (C) The primer support is then reacted with a phosphitylation reagent (amidite), which is dissolved in acetonitrile, the amidite having the formula: wherein bg is a 5′-blocking group, 1 g is a leaving group, G is O or S, pg is a phosphorus protecting group, and R2′ and Bx have, independent of the analogous variables on the primer support, the same definitions as previously defined.        
The product of this reaction is the support-bound phosphite dimer: 
Support-bound wherein each of the variables bg, pg, G, R2′ and Bx is independently defined above, link is the linker and supt is the support, as defined above.                (D) The support-bound dimer is then typically washed with acetonitrile.        (E) A capping reagent in acetonitrile is then added to the column, thereby capping unreacted nucleoside.        (F) The column is then washed again with acetonitrile.        (G) The support-bound dimer is then typically reacted with an oxidizing agent, such as a thiolating agent (e.g. phenylacetyl disulfide), in acetonitrile, to form a support-bound phosphate triester: wherein G′ is O or S and the other variables are defined herein.        (H) The support-bound phosphate triester is then typically washed with acetonitrile.        
Steps (A)–(F) are then repeated, if necessary, a sufficient number of times to prepare a support-bound, blocked oligonucleotide having the formula: wherein n is a positive integer (typically about 7 to about 79).
The phosphorus protecting groups pg are then typically removed from the oligomer to produce a support-bound oligomer having the formula: which, after washing with a suitable wash solvent, such as acetonitrile, is typically cleaved from the solid support, purified, 5′-deblocked, and further processed to produce an oligomer of the formula: 
The person having skill in the art will recognize that G′H bound to a P(V) phosphorus is generally is ionized at physiologic pH, and that therefore, wherever G′H appears in the formulae above, or hereafter, G′− is synonymous therewith (the O− or S− being countered by a suitable cation, such as Na+).
In the foregoing background, bg is a blocking group, such as an acid-labile group. Such groups include the monomethoxytrityl group (MMT), the dimethoxytrityl group (DMT), the pixyl group, etc. DMT is the most widely used 5′-hydroxylprotection group for nucleoside derivatives used in automated nucleic acid synthesizers. DMT is also known in the literature as 4,4′-dimethoxytriphenylmethyl (and is alternatively abbreviated DMTr). DMT has the advantage of being nearly quantitatively removed with dilute acid during cyclical automated synthesis. The removed DMT is in the form of a cation, which can be detected and measured by an in-line ultraviolet spectrophotometer to indicate the progress of the reaction. During the synthesis of the nucleoside monomers, the DMT group is selective for the 5′ primary hydroxyl over the 3′ secondary hydroxyl of the nucleoside. Typically, the reaction between the base-protected nucleoside and DMT chloride is carried out in pyridine as a solvent/base with 1.2–1.3 molar equivalents of the DMT chloride. The selectivity for the 5′-OH is not optimal, so initially there is a mixture of 5′-O-mono and 3′-O-mono substituted DMT products. As the reaction completes it is desirable to force the reaction to the point in which the 3′ mono DMT is converted to the 3′,5′ bis DMT product because it is easier to separate. The mono DMT derivatives have very similar physical properties and are difficult to separate. Furthermore, the 3′ DMT product will react in the next step, phosphitylation, and will not be separable and thus will be incorporated into the desired oligonucleotide. It will also be difficult to detect once incorporated as the mass of the resulting oligonucleotide is the same. In the process, the 5′ mono desired product is also partially converted to the bis product. As a result, the yield of purified 5′ mono DMT product is reduced substantially. Typically yields of 70–80% of theory are the best to be expected. This reduction in yield is especially harmful when using more expensive, modified nucleosides such as 2′-O-alkyl-ribonucleosides.
There is a need for a method of regioselectively protecting a nucleoside, especially at the 5′-position of a ribonucleoside or a deoxy ribonucleoside, which provides for yields of protected nucleoside of greater than about 80% of theory. There is especially a need for such a method that provides for yields of greater than about 85% of theory, preferably of greater than about 90% of theory, and more preferably of greater than about 95% of theory.
There is also a need for a method of regioselectively protecting a nucleoside, which provides a nucleoside having excellent purity.
There is also a need for a method of regioselectively protecting a nucleoside, which provides greater regioselectivity than pyridine in the analogous reaction.
There is also a need for a protected nucleoside, especially a 5′-protected nucleoside, which has a minimal degree of 3′-protected impurity. There is especially a need for a 5′-protected nucleoside that has less than about 1%, and more preferably less than about 0.5%, and even more preferably less than about 0.1% of the 3′-protected impurity.
There is also a need for a method that provides a 5′-protected nucleoside that is suitable for preparing phosphoramidites for use in automated synthesis of oligonucleotides, which method excludes a chromatography step.
There is also a need for a method of preparing a 5′-protected phosphoramidite, wherein the phosphoramidite is substantially free of the 3′-protected impurity.